Diminished immune cell adhesion in hypoimmune ICAM-1 knockout human pluripotent stem cells

Saha, Sayandeep; Haynes, W. John; Seo, Jiwon; Del Rio, Natalia M.; Young, Elizabeth E.; Zhang, Jue; Holm, Alexis M.; Pimentel, Mireya; Flannagan, Lauryn; Huang, Liupei; Blashka, Wesley; Murphy, Lydia; Scholz, Merrick J.; Henrichs, Abigale; Suresh Babu, Jayalaxmi; Steill, John; Kratz, Jeremy; Stewart, Ron; Kamp, Timothy J.; Brown, Matthew E.

doi:10.1038/s41467-025-62568-2

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Published: 12 August 2025

Diminished immune cell adhesion in hypoimmune ICAM-1 knockout human pluripotent stem cells

Nature Communications volume 16, Article number: 7415 (2025) Cite this article

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Abstract

Gene edited human pluripotent stem cells are a promising platform for developing reparative cellular therapies that evade immune rejection. Existing first-generation hypoimmune strategies have used CRISPR/Cas9 editing to modulate genes associated with adaptive immune responses, but have largely not addressed the innate immune cells, such as neutrophils, that mediate inflammation and rejection processes occurring early after graft transplantation. We identify the adhesion molecule ICAM-1 as a hypoimmune target that plays multiple critical roles in both adaptive and innate immune responses post-transplantation. In our experiments, we find that ICAM-1 blocking or knockout in human pluripotent stem cell-derived cardiovascular therapies imparts significantly diminished binding of multiple immune cell types. ICAM-1 knockout results in diminished T cell proliferation and activation responses in vitro and in longer in vivo retention/protection of knockout grafts following immune cell encounter in NeoThy humanized mice. We also introduce the ICAM-1 knockout edit into existing first-generation hypoimmune human pluripotent stem cells and prevent immune cell binding. This promising hypoimmune editing strategy has the potential to improve transplantation outcomes for regenerative therapies in the setting of cardiovascular pathologies and several other diseases.

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Introduction

Human pluripotent stem cell (hPSC)^1,2 technology is a promising cellular therapy platform for treating multiple diseases, including acute cardiovascular pathologies (e.g., myocardial infarction)³, Parkinson’s disease⁴, and autoimmune disorders (e.g., type 1 diabetes)⁵. Two key attributes of hPSCs are: 1) scalability, which allows cells from a single donor to treat potentially thousands of patients; and 2) amenability to gene editing, including hypoimmune strategies to modulate genes involved in alloimmune rejection with the goal of achieving immune tolerance and long-term retention of hPSC-based grafts⁶. The first generation of hypoimmune hPSC gene edits largely focuses on targeting important mechanisms of adaptive immunity-driven allograft rejection, namely T cell-mediated rejection and antibody-mediated rejection. Hypoimmune editing strategies include genetic knockout (KO) of major histocompatibility complex (MHC) class I and II surface expression^7,8, individually and with additional knock-in edits to mitigate natural killer (NK) cell-mediated cytotoxicity (an adverse effect resulting from MHC KO)^9,10. While these adaptive immunity-centric methods are promising, innate immune cells (e.g., neutrophils, monocytes) are intimately involved in the very earliest stages of allograft rejection of solid organs and cellular therapies^{11,12,13,14,15} and thus remain a critical concern that has not been fully addressed by existing hypoimmune strategies¹⁶.

We sought to identify clinically-relevant hypoimmune editing targets by conducting literature searches on genes involved in adaptive as well as innate immune allograft rejection pathways, prioritizing 1) those that were utilized by multiple immune cell types and 2) those where the literature showed viable KO mice could be created (i.e., implying that the KO’d gene is not essential for survival). Intercellular Adhesion Molecule 1 (ICAM-1) is a glycoprotein expressed on the cell surface and in secreted forms by a variety of cells including endothelial cells (ECs), cardiomyocytes (CMs), epithelial cells, fibroblasts, and various immune cells. Adaptive and innate immune cells express ICAM-1 binding partners: the integrins LFA-1 (CD11a/CD18) and/or MAC-1 (CD11b/CD18) (Fig. 1a, and Supplementary Fig. 1). Immune cell LFA-1 and/or MAC-1-mediated binding to ICAM-1 on ECs is a critical step in the very earliest inflammatory stages of transplant allograft rejection, as it stabilizes the immune synapse that precedes effector-mediated killing and plays a key role in the firm adhesion process required for extravasation of immune cells into the parenchyma of transplanted grafts¹⁷. These binding interactions can also directly impact T cell activation¹⁸ and effector function¹⁹, making ICAM-1 an attractive editing target with potentially pleiotropic hypoimmune effects. Previous studies in mice have demonstrated that KO of ICAM-1 isoforms may have a beneficial impact on oxidative stress responses during inflammation²⁰ and that absence of surface ICAM-1 diminishes immune cell binding to ECs²¹ and other parenchymal cell types, resulting in prolongation of allograft survival in heart transplant studies²². Clinical studies have assessed targeting ICAM-1 via blocking antibodies^23,24,25 and antisense oligonucleotides²⁶ in transplantation and other settings, with conflicting results. The systemic nature of these approaches and/or recombinant antibody design limitations in these early trials may have obscured the potential of anti-ICAM-1 interventions. Nonetheless, clinical investigation with advanced ICAM-1 blocking antibodies is still an active area of research in the human solid organ²⁷ and xenotransplantation²⁸ fields.

**Fig. 1: Adhesion Assay with Antibody Blocking of ICAM-1 in Pluripotent Stem Cell (PSC)-Derived Cardiovascular Cells.**

The advent of CRISPR/Cas9-based gene editing^29,30 has enabled design of targeted (i.e., cell-specific) gene edits for personalized hPSC-based cellular therapies, potentially overcoming some of the barriers that hampered previous attempts to leverage systemic blockade of ICAM-1 for immune tolerance induction. To assess the potential of ICAM-1 as a hypoimmune target in hPSCs, rather than just in primary cell types, we used ICAM-1 blocking antibodies in in vitro cultures of clinically relevant hPSC-based cardiovascular therapies (hPSC-CVTs; including CMs, ECs, and cardiac fibroblasts [cFibs]) to verify inhibition of immune cell binding. We also used a targeted CRISPR/Cas9 editing strategy to create hPSC lines that were null for surface and secreted ICAM-1 expression. ICAM-1 ablation dramatically diminished binding of multiple immune cell types to hPSC-CVTs, and impeded in vitro and in vivo immune responses to the grafts. This hypoimmune strategy was effective at diminishing immune cell adhesion as a stand-alone gene edit, and when added as a secondary KO to existing first generation hypoimmune MHC KO hPSC lines.

Results

ICAM-1 blockade reduces leukocyte adhesion to hPSC-derived cellular therapies

To validate the overall approach of targeting ICAM-1 in hPSCs to promote immune evasion, we utilized established ICAM-1 blocking antibodies for classical in vitro binding assays. Multicellular hPSC-CVTs are a promising strategy for generating mature, vascularized grafts to repair damaged heart tissue following myocardial infarction^31,32. We focused our in vitro studies on three hPSC-derived cell types commonly utilized in these grafts: ECs, CMs, and cFibs. ECs were selected as the initial cell type to test the blocking strategy, as they are the first target of allograft rejection in CVTs and other vascularized hPSC-based therapies, as well as in cardiac allograft vasculopathy and other adverse events that occur during rejection of traditional solid organ transplant grafts^33,34. The H9 human embryonic stem cell line was differentiated into highly pure ( >90% CD31⁺CD144⁺) ECs using a standardized seven day protocol³⁵, then cultured and stimulated for 48 h with pro-inflammatory cytokines tumor necrosis factor alpha (TNFα; 10 ng/ml) and interferon gamma (IFNγ; 50 ng/ml) to mimic the early inflammatory post-transplantation microenvironment. Treating H9 ECs with anti-ICAM-1 blocking antibody (0.5 mg/µl) for 1 h at 37 °C resulted in significantly diminished binding of the monocytic U937 cell line to the ECs (Fig. 1b). ICAM-1 blocking also impeded U937 binding in hPSC-derived CMs (Fig. 1c) and cFibs (Fig. 1d). U937 cells-only controls did not bind to Matrigel-coated wells after being washed similarly to experimental co-culture wells. These results implicate ICAM-1 as a critical molecule for immune cell binding to multiple PSC-derived cell types, and are consistent with the literature investigating primary (non-PSC) cell types from humans^27,36, mice²¹, and other species³⁷.

Generation of hypoimmune ICAM-1 KO hPSCs

To generate hypoimmune PSCs, we employed a CRISPR/Cas9-based strategy to KO ICAM-1 in multiple hPSC lines via introduction of a stop codon, preventing generation of any isoform of the ICAM-1 protein (Fig. 2a). ICAM-1 KO hPSCs were indistinguishable from isogenic wild-type (WT) hPSCs by each of the several hPSC characterization metrics we tested. KO lines showed typical hPSC morphology (Fig. 2b). They had similar pluripotency compared to WT, as assessed by staining for SSEA-4 and alkaline phosphatase, and in teratoma assays (Fig. 2c, d and Supplementary Fig. 2a). Cells were karyotypically normal after gene editing as determined by g-banding method (Supplementary Fig. 2b), and no off-target effects of CRISPR/Cas9 editing were detected via Sanger sequencing of the five highest-likelihood off-target sites predicted by CRISPOR algorithms (see Data Availability, below). WT and KO lines both displayed normal responses to inflammatory stimulation (TNFα [10 ng/ml] and IFNγ [50 ng/ml] for 48 h) by upregulating expression of MHC class I on their cell surfaces (Fig. 3a). As expected, WT hPSCs showed baseline expression and upregulation of surface ICAM-1 following inflammatory stimulation, but in the ICAM-1 KO lines no protein was detectable by flow cytometry (Fig. 3b; and Supplementary Fig. 1) or Western Blot (Fig. 3c). Lastly, we assessed global transcriptional profiles as well as expression of pluripotency-associated genes by bulk RNA sequencing and found no meaningful differences between WT and KO hPSCs (Fig. 3d, e). Our results indicate that our gene editing was effective in ablating functional ICAM-1 protein (i.e., a complete KO) in all lines tested, that the integrity and stability of the hPSC lines is maintained, and that ICAM-1 KO hPSCs are suitable for subsequent differentiation into cellular therapy candidates and immunogenicity studies.

**Fig. 2: Generation of ICAM-1 Knockout Pluripotent Stem Cell (PSC) Lines.**

**Fig. 3: Transcriptomic and Protein Expression Analysis of ICAM-1 Knockout (KO) vs Wild Type (WT) Cells.**

ICAM-1 KO hPSCs have typical differentiation capability

An advantage of hypoimmune gene editing is the theoretical possibility of generating one universal donor hPSC line that can be thoroughly validated for safety and other attributes prior to widespread clinical use in large cohorts of patients. This is in contrast with autologous hPSCs, where each newly-created patient-derived line will likely require batch-specific testing and regulatory approval prior to use³⁸. Given the potential use of hypoimmune hPSCs in multiple disease and clinical contexts, it is critical to assess their capability to differentiate into multiple lineages and validate each universal line’s utility for each disease-specific therapeutic application. As shown in Fig. 2, ICAM-1 KO hPSCs formed teratomas and spontaneously differentiated into cell types from each of the three germ layers, indicating robust pluripotency potential. We next assessed the capability of ICAM-1 KO cells to be directed to differentiate into specific hPSC-CVT cell types using well-established protocols^35,39,40. Similar to their isogenic WT hPSC controls, ICAM-1 KO cell lines differentiated into high-purity CD31⁺CD144⁺ ECs, cardiac troponin T (cTNT)⁺ CMs, and TE-7⁺ cFibs (Fig. 4a–c). No evidence of SSEA-4⁺ undifferentiated hPSCs were seen in EC, CM, or cFib in vitro cultures, and upon injection of hPSC-CVTs into immune-compromised mice, teratomas were only observed in animals receiving hPSC control injections (Supplementary Fig. 3). These results show the potential use of ICAM-1 KO hPSCs as a universal cell line for generation of reparative CVTs. We next needed to assess immune cell interactions, as this metric is critical for determining their clinical potential.

**Fig. 4: ICAM-1 Knockout (KO) Pluripotent Stem Cell (PSC) Differentiation Capacity and Surface ICAM-1 Protein Expression.**

Surface and soluble ICAM-1 expression is ablated on differentiated cells

In humans and mice, there are multiple isoforms of ICAM-1 generated via alternative splicing and proteolytic cleavage events that occur during inflammatory responses. These isoforms are expressed at the cell surface and in a secreted soluble form (sICAM-1), so we assessed our hypoimmune hPSCs for both. Similar to the parent hPSCs, there was no surface ICAM-1 detected in any of the hPSC-differentiated cell types by flow cytometry (Fig. 4d), indicating that the KO was thorough, effective, and devoid of the leakiness that has been documented in some previous hypoimmune gene editing approaches (e.g., MHC class I via KO of beta-2 microglobulin [β2M])^10,41. The differentiated cells also maintained normal MHC class I upregulation responses to inflammatory stimuli (Supplementary Fig. 4).

Our gene editing strategy introduces a stop codon in the first exon of the ICAM-1 gene on Chromosome 19, preventing translation and subsequent formation of any complete isoform protein. No surface protein was detectable in any of the independently-generated ICAM-1 KO hPSC lines made by our team (n = 6). To validate KO of sICAM-1 (i.e., complete nullification of all ICAM-1 protein products), we performed a Luminex assay on cell culture supernatants from hPSC-ECs that had been stimulated, as described in Figs. 1 and 4 above, for 48 h with inflammatory cytokines. WT hPSC-ECs secreted large quantities ( > 100 ng/ml) into cell culture supernatants following stimulation, while KO cells have no detectable levels (Fig. 5). These results confirm the effectiveness of our KO strategy for total ablation of surface as well as secreted isoforms of ICAM-1.

Immune cell adhesion is diminished in ICAM-1 KO cells

Immune cell binding to ECs (as well as to CMs, cFibs, and other parenchymal cells) is stabilized via ICAM-1 and is a critical first process in the early, inflammatory stages of allograft rejection following solid organ and cellular therapy transplantation. To assess this immune cell binding, we first used the well-established U937 monocytic lymphoma cell line, which expresses the ICAM-1 binding ligands LFA-1 and MAC-1 (see Fig. 1a), to model early innate immune cell interactions with hPSC-ECs following transplantation. High purity hPSC-ECs were differentiated from WT and ICAM-1 KO lines, stimulated for 48 h with TNFα (10 ng/ml) and IFNγ (50 ng/ml), then co-cultured as described in Fig. 1 with fluorescently labeled U937 cells. ICAM-1 KO ECs showed dramatically diminished binding of U937s compared to WT controls (Fig. 6a; and Supplementary Fig. 5a).

**Fig. 6: Immune Cell Adhesion Assay with ICAM-1 Knockout (KO) Pluripotent Stem Cell (PSC)-Derived Endothelial Cells (ECs).**

Neutrophils are another primary responder of the innate immune system and they play a significant role in the rejection of solid organ transplants. Understanding the multifaceted role of neutrophils in transplant rejection and experimentally testing the neutrophil binding response to hPSC-derived grafts is crucial for mitigating their detrimental effects and prolonging graft survival^42,43. Similar to the U937 binding experiments described above, we fluorescently labeled allogeneic neutrophils and co-cultured them with hPSC-ECs, showing diminished binding to ICAM-1 KO cells (Fig. 6b; and Supplementary Fig. 5b). We also performed binding experiments with peripheral blood mononuclear cells (PBMCs; i.e., a mixture of innate and adaptive LFA-1⁺ and MAC-1⁺ cells, but lacking neutrophils or other granulocytes) from n = 2 human leukocyte antigen (HLA)-mismatched donors (Fig. 6c; and Supplementary Fig. 5b, Supplementary Table 1). All innate and adaptive immune cells tested in this series of experiments showed significantly diminished binding to hPSC-derived cells, indicating a broad anti-adhesion effect of ICAM-1 KO.

ICAM-1 KO is protective against in vitro and in vivo immune responses

ICAM-1 KO, similar to antibody blocking, dramatically reduces but does not completely prevent binding of innate and adaptive immune cells to hPSC-derived cells. We hypothesized that the very few numbers of immune cells that do attach to hPSC-ECs would have attenuated immune responses to the other ICAM-1 KO cell types within the hPSC-CVT. We conducted a version of the mixed lymphocyte reaction (MLR)⁴⁴ to assess the alloantigen-induced proliferative responses of HLA-mismatched T cells to the hPSC-CVTs (i.e., to model the effect of cells that bound ECs and were able to extravasate into the graft, then encountering ICAM-1 KO parenchymal cells). In response to 6 day MLR co-culture with hPSC-ECs and -CMs, we observed significantly diminished proliferative responses of total CD8 + T cells, as well as the CD8+ effector memory (T_EM) CD45RO⁺ CD45RA^- CD62L^- CCR7^- cell subset known to play a prominent role in T cell-mediated rejection⁴⁵ (Supplementary Fig. 6a). Importantly, there was a significant reduction in both proliferation and activation of T cells (Fig. 7a, b), highlighting the role of ICAM-1 in multiple aspects of T cell effector function. These data indicate that the lack of ICAM-1 binding resulted in a diminished proliferation response in this alloreactive T cell population.

**Fig. 7: In Vitro and In Vivo Immune Responses to ICAM-1 Knockout (KO) Cells.**

We then tested in vivo immune responses to hypoimmune cells using the NeoThy, an advanced human immune system humanized mouse model created by intravenous injection of neonatal human cord blood hematopoietic stem and progenitors and surgical implantation of human thymus tissues^46,47. Similar to the fetal tissue-based bone marrow-liver-thymus (BLT) mouse model, NeoThy mice engraft with a robust repertoire of adaptive and innate immune cells but use more developmentally-mature non-fetal tissue sources. We transplanted WT and ICAM-1 KO hPSCs expressing constitutive Akaluc bioluminescence imaging (BLI) reporter constructs, co-injected with Matrigel, into the hind limbs of NeoThy mice with verified human immune chimerism (made with HLA-mismatched humanizing tissues) exceeding 10% human CD45⁺ and 10% human CD3⁺ cells in circulation. In each individual humanized recipient (n = 5), WT cells were injected into the left mouse leg and KO cells into the right leg (Fig. 7c; and Supplementary Fig. 6b). In 4 of 5 mice, KO cells were retained and grew into teratomas, similar to non-humanized controls used for the teratoma assays in Fig. 2d. In contrast, WT cells were controlled or diminished in size, as evidenced by live animal imaging of BLI signal over three timepoints up to 32 days post-transplant. Taken together, the in vitro and in vivo immune assay results indicate an overall differential and more robust immune response to WT cells compared to KO cells. The previous binding assay data offer additional evidence implicating immune cell binding, or lack thereof, as a critical mechanism mediating these results. Overall, these findings support the use of ICAM-1 KO hPSCs as a hypoimmune therapy platform with potential to elicit significantly attenuated allograft rejection responses following transplantation into patients.

ICAM-1 KO confers additional protection to first-generation hypoimmune hPSCs

Having demonstrated that ICAM-1 KO diminishes immune cell binding as a stand-alone edit, we sought to validate addition of ICAM-1 KO to a promising, previously published first generation hypoimmune hPSC line to test whether it was possible to confer anti-adhesion properties to the line. We first engineered the H9 human embryonic stem cell PSC line with a constitutively expressed Akaluc GFP construct to enable BLI⁴⁸, followed by a double (β2M and CIITA) KO coupled with overexpression of NK cell inhibitory ligand CD47, as described by Deuse et al.⁹. This line was called H9 DKO. We then added ICAM-1 KO to H9 DKO, as described for editing of WT lines above, to make the H9 TKO line (Supplementary Fig. 7a–c). As expected, after differentiation into highly pure ECs, both H9 DKO and H9 TKO did not express or upregulate MHC class I after stimulation for 48 h with TNFα (10 ng/ml) and IFNγ (50 ng/ml). Cell surface ICAM-1 and sICAM-1 expression at baseline and following stimulation was still present in H9 DKO, indicating that ICAM-1-mediated immune cell binding and associated allograft rejection events are still an active concern for first generation hypoimmune hPSCs. As expected, ICAM-1 KO ablated all isoforms of ICAM-1 in H9 TKO (Fig. 8a–c; and Supplementary Fig. 7 d). Concordantly, the ICAM-1 null H9 TKO line-derived ECs elicited diminished binding of U937 cells and PBMCs (Fig. 8d) following stimulation in leukocyte adhesion assays. CD8⁺ T cell activation and proliferation were also significantly diminished in the H9 TKO-ECs compared to H9 WT cells (Fig. 8e).

**Fig. 8: Addition of ICAM-1 Knockout (KO) to First Generation Hypoimmune Pluripotent Stem Cell (PSCs).**

Discussion

Here, we showed that ICAM-1 KO has clear effects in diminishing binding of innate and adaptive immune cells and that evasion of in vitro and in vivo immune recognition is conferred by this editing approach. There are a variety of promising hypoimmune gene editing strategies to prevent adaptive immune responses to hPSC-derived grafts^7,9,10,49, but to date no strategy has been demonstrated to blunt all components (cellular and humoral) of the multifaceted allograft rejection response. In this study, we leveraged two critical technological advances (hPSCs and CRISPR/Cas9 gene editing) to precisely introduce ICAM-1 KO into hPSC-CVT cell types. Similar to xenotransplantation gene editing strategies⁵⁰, it may be useful to KO multiple genes in a single universal hypoimmune hPSC line to promote evasion of the coordinated multicellular immune responses that occur in vivo. Such strategies will need to be validated in multiple hPSC-based cell therapies and in several disease contexts, and certain disease-specific therapies may need more editing than others. Additionally, introduction of multiple gene edits in one cell line will need to balance effectiveness with avoidance of off-target effects and other adverse events^51,52.

Interestingly, in the late 1990s/early 2000s researchers did not yet know of ICAM-1 alternative splicing and multiple groups generated ICAM-1-deficient mouse strains that were not completely null for all functional isoforms. Indeed, as described by Ramos et al., the majority of all existing literature describing ICAM-1 KO mice does not include true ICAM-1 KOs, but rather mice only deficient in specific isoforms⁵³. While ICAM-1 KO mice have since been generated⁵⁴, the effect of this significant oversight still impacts the knowledge base in this area. As sICAM-1, which can be isoform-associated, has been used as a biomarker for a number of inflammation-associated pathologies, including spontaneous and transplantation-associated coronary artery disease⁵⁵, the impact of its absence in future hypoimmune hPSC therapies may have relevance for tissue repair following transplantation in terms of both biological function and clinical assessment of graft failure. We demonstrated that addition of ICAM-1 KO to first generationβ2M and CIITA double KO/CD47-overexpressing cells results in karyotypically normal hPSCs with diminished immune cell-binding, proliferation, and activation propensity, in addition to retaining the original hypoimmune benefits of the parent line. Future studies will be needed to discern whether stand-alone ICAM-1 KO is sufficient for all clinical applications or whether combining ICAM-1 KO with other strategies is ideal. Additionally, while we showed no evidence of undifferentiated hPSCs in our cell product, prior to initiation of clinical trials any hPSC-based therapy will need to undergo rigorous safety testing to evaluate propensity for malignancy, viral infection, and teratoma formation. This is especially true of all types of hypoimmune therapies, as they may escape mechanisms that the immune system uses to prune aberrant cells⁵⁶.

During the preparation of this manuscript, another research group published encouraging findings with introduction of ICAM-1 KO along with CD58 KO into first-generation MHC KO PSC lines in the context of CAR NK cell therapies for cancer, with the goal of preventing NK cell-mediated missing-self killing⁵⁷. These results corroborate our findings (i.e., ICAM-1 KO impairs cell adhesion) in another important research context. Additional studies will be needed to assess whether prevention of NK cell missing-self killing is a significant concern when MHC I and II are intact, as is the case with our ICAM-1 KO hPSCs. ICAM-1 participates in T cell activation, and our in vitro T cell studies showed that ICAM-1 KO directly impacts T cell alloresponses; hypoimmune hPSCs may therefore not need MHC KO to mitigate T cell-mediated rejection. Antibody-mediated rejection, however, is an important driver of allograft rejection and MHC I and II are primary targets for donor-specific antibodies, so MHC KO combined with ICAM-1 KO may still be advantageous for transplanting into highly-sensitized patients⁵⁸. Careful crossmatch assessment prior to transplantation could potentially minimize the incidence of antibody-mediated rejection in hPSC-CVT transplantation^59,60. It should be noted that antibody-mediated rejection can involve antibodies targeting a variety of non-HLA epitopes (e.g., EC-specific surface molecules)^61,62,63, so it may not be possible to create a universal cell line via MHC ablation alone that is completely effective in eliminating this powerful driver of allograft rejection. Other additive hypoimmune editing approaches may be warranted, such as a CD64 overexpression strategy recently described by Gravina et al.⁴⁹. Relatedly, better in vitro and in vivo models of human antibody-mediated rejection are needed to adequately interrogate these types of research questions and additional studies assessing impact of donor:recipient HLA disparities may be insightful. It is important to note that donor-specific antibodies play an important role in the process of complement fixation during allograft rejection, and complement binding is known to upregulate ICAM-1 expression on endothelial cells, which in turn initiates deleterious immune cell adhesion-mediated events^64,65. While our ICAM-1 KO approach is not intended to block complement or antibody binding directly, this KO is likely to interrupt the pathological immune adhesion portion of cascades that result from such binding, further favoring the development of immune tolerance.

Humanized mouse models are quite useful for T cell-mediated rejection and NK cell-mediated allograft rejection studies^66,67,68,69, but the most commonly used humanized mice are suboptimal for antibody-mediated rejection studies due to B cell immaturity and impeded class switching in de novo antibodies⁷⁰. Our group and others are working on next-generation iterations of humanized mouse models that incorporate more robust T cell, NK cell, and innate immune cell repertoires in order to improve their utility for transplantation immunology-focused research studies, and creation of tractable models harboring antigen-specific IgG antibody pools is an area of intense focus. We anticipate that improved next-generation humanized mouse models will become a critical tool in the clinical translation of advanced hypoimmune gene edited hPSC therapies in the coming years.

Future applications of ICAM-1 KO and related editing could include optimization of engineering strategies to attenuate, but not totally ablate, immune responses to balance diminishing allograft rejection and graft loss with allowing for some amount of residual immunosurveillance capacity against malignant transformation and/or formation of viral (e.g., cytomegalovirus) reservoirs. Additionally, our ICAM-1 KO developed for use in hPSC therapeutics could also be a promising target for gene editing in xenotransplantation. ICAM-1 is highly conserved across species, and previous studies have shown that human cells bind to pig cells via ICAM-1⁷¹. Lastly, ICAM-1 is the primary receptor for human rhinovirus⁷² and also plays a role in cell-to-cell transmission of human immunodeficiency virus⁷³. We intend to explore the impact of ICAM-1 KO on viral infection dynamics in future studies.

Our study highlights the importance of designing hypoimmune gene editing approaches with adaptive and innate immune cells in mind. Targeting ICAM-1 as a common molecule utilized by multiple cell types during the earliest steps in the allograft rejection process demonstrates a new paradigm for developing clinically-relevant hypoimmune gene edits. Further progress in this area of research will ultimately improve the effective clinical translation of regenerative therapies for cardiovascular pathologies and other diseases.

Methods

This work was approved by the Animal Care and Use Committee of the University of Wisconsin (UW) School of Medicine and Public Health and the UW Health Sciences Institutional Review Board, and complied with federal and state law.

Genetic modification of PSC lines

KO of ICAM-1

Multiple human induced PSC (PED05, XY, derived in our lab; 19-9-11 [i.e., iPS-DF19-9-11T], XY, from WiCell) and embryonic stem cell lines (H9/WA09, XX; H1/WA01, XY, both from WiCell) were edited. The single guide RNA (sgRNA, CCCGCACTCCTGGTCCTGCTCGG) for the site of interest was identified using the CRISPOR design tool⁷⁴. 1.5 nmol synthetic sgRNAs were constructed with 2’-O-methyl 3’ phosphorothioate modifications at the first and last 3 nucleotides (Synthego). In preparation for electroporation, PSCs were cultured on Matrigel in TeSR-PLUS media (StemCell Technologies) and grown to ~80% confluency following standard cell culture protocols. Cells were treated with CloneR (StemCell Technologies) 24 h before electroporation per the manufacturer’s protocol. Preceding electroporation, reconstitution of sgRNA constructs to a concentration of 150 pmol/mL was conducted following the manufacturer’s protocols. We combined 4 mg of Cas9 Nuclease protein (TrueCut Cas9 Protein V2, Thermo Fisher Scientific), 5 mL of Neon Buffer R (Invitrogen), and 1 mL of reconstituted sgRNA to promote Cas9- ribonucleoprotein (RNP) complex formation. After an incubation of 15 min, 1.0 mL of ssODN primer designed with homology overhangs of at least 40 base pairs (reconstituted to 1 mg/mL) was added into the Cas9-RNP mixture. PSCs were detached and singularized utilizing a 1:1 mixture of 0.5 mM EDTA and Accutase (Invitrogen) via incubation with EDTA/Accutase for 3–4 min, resuspension in 1 mL phosphate buffered saline (PBS; Corning), and centrifugation into a pellet. The cells were resuspended in 35 mL Buffer R (Thermo Fisher Scientific) and combined with 8 mL of the previously prepared Cas9-RNP complex with repair ssODN. Cells underwent electroporation via 10 μL NEON electroporation format with 1x pulse, 30 msec, and 1200 V settings. Following four subsequent rounds of electroporation, cells were pooled and plated in TeSR-PLUS (StemCell Technologies) maintenance media supplemented with CloneR (StemCell Technologies) per the manufacturer-recommended concentrations utilizing serial dilutions to foster single-cell clonal growth. After expansion over a span of 10-14 days, clones were identified and picked using standard techniques. QuickExtract DNA Extraction Solution 1.0 (Epicenter) was used to generate bulk gDNA from dissociated cells and a portion of clones to validate editing efficiency before clonal selection. Single-cell clones were manually selected and subsequently mechanically disaggregated. Genotyping primers were created by designing flanks around the mutation site, promoting amplification of this region via Q5 polymerase-based PCR (New England Biolabs). PCR products were then identified by agarose gel method and purified via Zymoclean Gel DNA Recovery Kit (Zymo Research). Sanger sequencing of clones was performed by Quintara Biosciences to identify clones that displayed the proper genetic modification. Suspected off-target sites for genome modification were analyzed to identify if the CRISPR/Cas9 system had generated any non-specific genome editing. Genotyping primers were designed to amplify the 5 highest-likelihood off-target sites for each sgRNA as predicted by the CRISPOR algorithms. The resulting PCR products were identified by agarose gel method, purified via Zymoclean Gel DNA Recovery Kit (Zymo Research), and submitted for Sanger sequencing to Quintara Biosciences.

Knock-In of constitutive reporter constructs

Generation of base PSC lines with BLI and fluorescence reporters for downstream studies and additional editing (detailed below) was conducted as described in Zhang et al.⁴⁸. The H9 (WA09, XX) embryonic stem cell line (WiCell) and the PED05 (XY) induced PSC line derived in our lab⁴⁷ were edited for constitutive expression of the Akaluc bioluminescence reporter as well as green fluorescence protein (GFP) from Aequorea victoria⁷⁵. The 5’- and 3’-homology arms of the targeting vector were cloned into a drug resistance containing vector. gRNA was synthesized according to the manufacturer’s instructions (GeneArt™ Precision gRNA Synthesis Kit, Thermo Fisher Scientific). To achieve the best electroporation efficiency, hPSCs were passaged with EDTA (1:4 split) and cultured to reach 80–90% confluency 2 days before the experiment. On the day of the experiment, hPSCs were resuspended at a density of 1.25 x 10⁷ cells/mL in E8 medium supplemented with 10 μM Y27632. To prepare RNP, 0.6 µg gRNA was mixed with 2 µg Cas9 protein (TrueCut™ Cas9 Protein v2, Thermo Fisher Scientific) in 5 µl E8 medium supplemented with 10 μM Y27632. The mixture was incubated for 10 min at room temperature. Next, we added 5.5 µl cell suspension and 4 μg targeting vector to RNP. The electroporation was performed in the Neon Electroporation System (Thermo Fisher Scientific) with program 13. Cells were combined from 2–4 electroporations and seeded into one 10-cm dish coated with Matrigel. Drug selection was performed when cells reached 20% confluency. Surviving colonies were picked after drug selection and expanded in E8 medium. The edited lines were called H9 ALG and PED05 ALG, respectively, and used as WT (non-hypoimmune) controls for BLI studies.

Generation of human MHC double KO-CD47 overexpression cell lines

The H9 ALG line was also then edited using the Deuse et al. strategy⁹ to overexpress CD47 (NK cell inhibitory molecule), in addition to KO ofβ2M (to prevent MHC I surface expression) and CIITA (to prevent HLA class II gene expression and thus inhibit MHC II surface expression).β2M and CIITA gRNA were synthesized according to the manufacturer’s instructions (GeneArt™ Precision gRNA Synthesis Kit, Thermo Fisher Scientific). H9 ALG cells were resuspended at a density of 1.25 x 10⁷ cells/mL in E8 medium supplemented with 10 μM Y27632. To prepare RNP, 0.3 µg of each gRNA was mixed with 2 µg Cas9 protein (TrueCut™ Cas9 Protein v2, Thermo Fisher Scientific) in 5 µl E8 medium supplemented with 10 μM Y27632. The mixture was incubated for 10 min at room temperature. Next, a 5.5 µl cell suspension was added to RNP, and electroporation was performed in the Neon Electroporation System (Thermo Fisher Scientific) with program 16. To get single-cell clones without drug selection, 2.5% of the electroporated cells were seeded into one 10-cm dish coated with Matrigel. Single cell-derived clones were picked 1-2 weeks later without drug selection. This first generation hypoimmune line with bioluminescence reporter is referred to as H9 DKO. The H9 DKO line was further edited to KO ICAM-1 expression, as described above. The resulting line is referred to as H9 TKO.

Karyotyping

G-banding analysis was conducted by the WiCell Research Institute (Madison, WI) in accordance with a standardized protocol⁷⁶. Cultures were harvested following standard metaphase preparation procedures. Briefly, cultured cells were exposed to ethidium bromide as an intercalation agent followed by Colcemid (Thermo Fisher Scientific), which arrests the cells in metaphase. Once the incubation is complete, the cells are exposed to potassium chloride followed by Carnoy’s fixative. Fixed cells are dropped onto slides in an environmental chamber, banded with trypsin, and stained with Leischman’s stain before imaging and analysis.

SSEA-4 flow cytometry

PSCs and PSC-CVTs cultured on Matrigel-coated (Corning) cell culture plates were harvested and stained for the presence of SSEA-4 (clone MC813-70, Thermo Fisher Scientific) to characterize pluripotency per the manufacturer’s instructions. Each well was aspirated of residual cell culture media and incubated for 7 min at 37 °C with 1 mL 1X TrypLE (Gibco) to detach the cells from the plate. After incubation, 1 mL of MACs Separation Buffer (Miltenyi) was added to each well as a staining buffer to collect and transfer the detached cells to a microcentrifuge tube. The cells were then centrifuged at 300 × g for 5 min. The supernatant was aspirated, and each tube was stained with 5 μL of SSEA4 antibody and incubated in the dark for 20 min at 4°C. After the incubation, the cells were centrifuged at 300 × g for 5 min and the supernatant was aspirated. Flow cytometry was performed on a Cytoflex Flow Cytometer (Beckman Colter) and analyzed using FlowJo Version 10 software.

Alkaline phosphatase assay

All PSC lines were grown to near-confluence on Matrigel-coated (Corning) cell culture plates. Cells were then stained with Vector Blue Alkaline Phosphatase Substrate Kit III (Vector Laboratories) according to the manufacturer’s protocol to assess pluripotency.

PSC culture

All PSC lines were cultured on Matrigel-coated (Corning) cell culture plates and fed daily with 2.5 mL TeSR E8 media (StemCell Technologies, WiCell Formulation). Upon reaching full confluency, cells were passaged to fresh Matrigel-coated cell culture plates. Each well was aspirated of residual cell culture media and rinsed with 2 mL PBS (without Calcium or Magnesium [-/-]; Corning). To detach cells from the plate for passaging, each well was incubated with 1 mL 5 mM EDTA solution made by adding 500 μL 0.5 M EDTA (Thermo Fisher Scientific) to 500 mL PBS-/- at room temperature for 7 min. After incubation, each well was aspirated and TeSR E8 media (StemCell Technologies, WiCell Formulation) was used to dislodge the remaining colonies still attached to the wells and transfer the cells into a fresh Matrigel-coated cell culture plate. Passage number was recorded, with each new passage adding to the cumulative total (i.e., passage number was not reset upon gene editing but continued onward from the input passage number at the start of the editing process).

Differentiation of cardiovascular cells from PSCs

EC differentiation

PSC-derived ECs were differentiated from all the PSC lines using the STEMdiff Endothelial Differentiation Kit (StemCell Technologies) according to the manufacturer’s protocol. Prior to differentiation, PSCs were maintained in TeSR E8 media on Matrigel-coated cell culture plates until they reached full confluency. Following 7 days of differentiation, ECs cultured on Matrigel-coated cell culture plates were aspirated of residual Endothelial Induction Media (StemCell Technologies) and incubated for 5 min at 37 °C with 1 mL room temperature Accutase (Gibco) to detach the cells from the plate. After incubation, the ECs were collected and magnetically enriched using the CD34 microbead kit and MACS columns (Miltenyi Biotec) according to the manufacturer’s protocol. The purified cells (purity determined as mentioned below) were then plated on Animal-Component-Free coated cell culture plates using Endothelial Expansion Media (StemCell Technologies) for expansion. To harvest cells for flow cytometry, each well was aspirated of media after 3 days of the first passage, and incubated with 1 ml of room temperature Accutase for 5 mins at 37 °C to detach the cells from the plate. After incubation, 1 mL of MACs Separation Buffer (Miltenyi) was added to each well as a staining buffer to collect and transfer the detached cells to a microcentrifuge tube. The cells were then centrifuged at 300 × g for 5 min. The supernatant was aspirated, and each tube was stained with 5 μL of APC-Cy7 Mouse Anti-Human CD31 (PECAM-1) antibody (clone WM59, BD Biosciences) and 5 μL of APC anti-human CD144 (VE-Cadherin) antibody (clone BV9, BioLegend) to assess EC purity. The cells were then incubated in the dark for 20 min at 4 °C. After the incubation, the cells were centrifuged at 300 × g for 5 min and the supernatant was aspirated. Flow cytometry was performed on a Cytoflex Flow Cytometer (Beckman Colter) and analyzed using FlowJo Version 10 software.

CM differentiation

PSC-derived CMs were differentiated from all the PSC lines implementing the Alternate GiWi protocol, using small molecule temporal modulation of the Wnt pathway, as described in Lian et al.⁴⁰. Prior to differentiation, PSCs were maintained in TeSR E8 media on Matrigel-coated cell culture plates until they reached full confluency. Following 17 days of differentiation, CMs cultured on Matrigel-coated cell culture plates were harvested and stained for Anti-Cardiac Troponin T (clone 1C11) Antibody (Abcam) and Goat anti-Mouse IgG (H + L) Cross-Adsorbed Secondary Antibody Alexa Fluor 647 (Thermo Fisher Scientific) to assess CM purity as described by Zhang et al³⁹. Flow cytometry was performed on a Cytoflex Flow Cytometer (Beckman Colter) and analyzed using FlowJo Version 10 software.

cFib differentiation

PSC-derived cFibs were differentiated from all the PSC lines as described in Zhang et al.³⁹. Prior to differentiation, PSCs were maintained in TeSR E8 media on Matrigel-coated cell culture plates until they reached full confluency. Following 20 days of differentiation, cFibs cultured on Matrigel-coated cell culture plates were harvested and stained for Anti-Fibroblast Antibody Clone TE-7 (Millipore Sigma) and Antibody to Sarcomeric Myosin MF20 (University of Iowa Developmental Studies Hybridoma Bank) to assess cFib purity as described by Zhang et al.³⁹. Flow cytometry was performed on a Cytoflex Flow Cytometer (Beckman Colter) and analyzed using FlowJo Version 10 software.

Fluorescent labeling of effector immune cells

Staining of the effector cells with Calcein-AM (Invitrogen) was performed in accordance to manufacturers recommendations. After counting the effector cells, the requisite number of cells required for the experiment was first aliquoted and washed with RPMI-1640 without phenol red (Gibco). After the wash, the cells were resuspended to 2 − 4 × 10⁶ cells/mL with warm (37 °C) RPMI-1640 without phenol red. The cells were added with 2.98 μL of 3 μM Calcein-AM stock solution (Invitrogen) per mL of media, mixed well with a serological pipette, and incubated in the dark at 37 °C for 30 min. Following incubation, the cells were centrifuged twice at 450 × g for 5 min to remove excess Calcein-AM and resuspended at 1 × 10⁶ cells/mL with warm RPMI-1640 without phenol red. Violet Proliferation Dye 450 (VPD450; BD Biosciences) was used as an alternative to Calcein-AM for some experiments where the cell harbored a green fluorescent protein. Effector cells were counted and the requisite number of cells for the experiment were washed twice using 1 X PBS-/- to remove any residual serum proteins. The washed cells were then resuspended at 10e6/ml of PBS-/-. 1ul of 1uM VPD450 stock was added to the cells per mL of PBS-/- and incubated for 15 min in the dark at 37 °C. After incubation, the cells were washed (centrifuged at 300 g for 5 min), and the cell pellet was resuspended in warm RPMI-1640 without phenol red with 10% fetal bovine serum at 1 × 10⁶ cells/mL.

Leukocyte adhesion assays

Adhesion assays of fluorescently-labeled immune effector cells with PSC-derived cells were performed using a combination of methods described by Zhang et al.⁷⁷ and Wilhelmsen et al.⁷⁸. All the PSC-derived cell types (ECs, CMs, and cFibs) were grown on a 24 well plate (Corning) and stimulated with IFNγ (50 ng/ml) and TNFα (10 ng/ml) 48 h prior to the experiment. After incubation, cell culture supernatants were aspirated and the wells were washed twice with RPMI-1640 with 10% bovine serum albumin (wash media) at room temperature. For experiments that incorporated ICAM-1 blocking antibody, following the wash, 500ul of fresh cell culture media for each respective cell type were combined with 0.5 mg/µl purified anti-human ICAM-1 blocking antibody (Clone HA58, BioLegend), added to the wells, and incubated at 37 °C for 1 h. The HA58 clone recognizes an epitope in the extracellular D1 domain of the ICAM-1 molecule, which is shared by all surface and secreted isoforms⁷⁹. Following incubation, the wells were washed once with room temperature wash media to remove excess unbound blocking antibody, followed by addition of U937s, PBMCs, or neutrophils (1e6/ml), and then incubated for 20–30 min. After incubation with immune cells, the wells were washed twice with cold wash media to remove excess unbound immune cells. The wells were imaged at baseline (without immune cells) and both before and after wash, with both phase contrast and fluorescent settings using Echo Revolve | R4 Microscope enabled with Echo PRO imaging software by Bioconvergence (BICO) to document the confluency and the auto-fluorescence, respectively, prior to the addition of the effector cells. A minimum of five randomly selected regions of interest were imaged for both phase contrast and fluorescence settings; the images were then analyzed with ImageJ software (version 1.54 g) to determine the mean adhered effector cells to the PSC-derived cell monolayer. The exposure settings were maintained across all images.

Luminex for soluble ICAM-1

Luminex assays detecting soluble ICAM-1 (sICAM1) were conducted using the MILLIPLEX Human Sepsis Magnetic Bead Panel 1 Kit (Millipore Sigma) according to the manufacturer’s protocol. Assays were run on the Luminex 200 Instrument System (Millipore Sigma) utilizing xPONENT running software. Results were analyzed using Belysa 1.1.0 analysis software.

Cell surface ICAM-1 detection

Individual cells were assessed by flow cytometry for surface ICAM-1 expression using the HA58 antibody clone mentioned above (see Leukocyte Adhesion Assays), conjugated to PE or APC fluorochromes (Biolegend). Flow cytometry was performed on a Cytoflex Flow Cytometer (Beckman Colter) and analyzed using FlowJo Version 10 software.

Teratoma assay

To assess the formation of teratoma by all the PSC lines (WT and gene-edited), two confluent wells of PSCs were harvested from a Matrigel-coated (Corning) 6-well cell culture plate. 1 − 2 × 10⁶ cells were combined with 50 μL Matrigel (Corning) and injected subcutaneously into the hind leg of female NBSGW mice (6–8 weeks old). Teratomas were permitted to form for 6-10 weeks after the initial cell injections. We also performed a follow-up teratoma assay to assess teratoma potential of PSC-CVTs. A total of 5x10⁵ PSC-CVTs and PSCs (control) were injected into the right and left hind limbs, respectively, of female mice (11–12 weeks old). Teratomas began forming in the PSC-injected limbs within 3–4 weeks post-injection and were monitored for 14 weeks to allow the teratoma to grow to ~1 cm × 1 cm in size. Animals were then sacrificed and the teratomas were collected for histology. Teratoma tissue and the tissue from the PSC-CVT injected hind limbs was submitted to the UW Department of Pathology and Laboratory Medicine’s Translational Research Initiative in Pathology Core for histological analysis and processing for hematoxylin and eosin staining. All animal work was conducted according to relevant national and international guidelines under the approval of the UW Institutional Animal Care and Use Committee, and maximum teratoma size of 1.5 cm diameter was not exceeded.

HLA typing

HLA typing was performed on both H9 and PED05 PSC lines and for PBMC3 and PBMC5 donors. The LinkSeq HLA-ABCDRDQDP + 384 typing kit was used⁴⁶.

U937 cell line

The U937 cell line was obtained from ATCC. It exhibits monocyte morphology and was originally derived from malignant cells obtained from the pleural effusion from a patient with histiocytic lymphoma⁸⁰. LFA-1 (CD11a/CD18) was assessed with a FITC anti-human antibody (clone m24, Biolegend), and MAC-1 (CD11b/CD18) with a Brilliant Violet 510 antibody (clone ICRF44, Biolegend), by flow cytometry.

PBMC and neutrophil isolation

PBMCs from donors PBMC3 and PBMC5 were obtained from leukophoresis (BioIVT). Leukopaks were processed with Lymphocyte Separation Medium (Cellgro). PBMCs were collected via centrifugation using SepMate™ PBMC Isolation Tubes (StemCell Technologies) per the manufacturer’s recommendations. Neutrophils from an unidentified donor were obtained from whole blood using EasySep™ Direct Human Neutrophil Isolation Kit (StemCell Technologies) following the manufacturer’s instructions. The Easy 50 EasySep™ Magnet (StemCell Technologies) was used to isolate the neutrophil population (verified by flow cytometry using an APC-cy7 antibody specific for human CD16 (clone 3G8, BD Biosciences) and a PE antibody for human CD66b (clone C10F5, BD Biosciences). LFA-1 and MAC-1 were assessed as described in U937 Cell Line, above.

RNA sequencing analysis

Library preparation, sequencing, quality control, alignment, and expression estimation were performed at the UW-Madison Biotechnology Center’s Bioinformatics Core Facility (Research Resource Identifier - RRID:SCR_017769; Madison, WI, United States). Specifically, Illumina TruSeq RNA libraries were constructed and 150 bp paired-end sequencing was performed using the Illumina Hi-Seq 2500 platform. Read quality was evaluated using FastQC⁸¹. Bioinformatic analysis of RNASeq reads adhered to ENCODE guidelines and best practices for RNASeq⁸². Briefly, alignment of adapter-trimmed (Skewer v0.2.2b)⁸³ 2 × 150 bp paired-end strand-specific Illumina reads to the GRCh38.p14 genome was achieved with the Spliced Transcripts Alignment to a Reference (STAR v2.5.3a) software⁸⁴, and a splice-junction aware aligner using Ensembl annotation⁸⁵. Expression estimation was conducted using RSEM v1.2.3.1 (RNASeq by Expectation Maximization)⁸⁶. To test for differential expression among individual group contrasts, expected read counts were used as input into edgeR v3.16.5⁸⁷. R Studio v4.4.1 was used for heatmap analysis. Significance of the negative-binomial test was adjusted with a Benjamini–Hochberg false discovery rate correction at the 5% level⁸⁸. Before statistical analysis with edgeR, independent filtering was performed, requiring a threshold of at least 1 read per million in two or more samples.

Mixed lymphocyte reaction

MLRs were performed similar to our published method⁸⁹ and as described in other hPSC studies⁹⁰. [It should be noted that this version of the MLR is different than a classical MLR, as originally described by Bach and Hirschhorn⁴⁴; however, the fundamental concept of co-culturing allogeneic cells and monitoring for immune cell proliferation and activation is similar]. Briefly, highly pure hPSC-derived cells ( > 99% CD31⁺CD144⁺ for ECs, > 85% cTNT⁺ for CMs) were derived from H9 WT and the gene-edited lines as mentioned in the methods above. The hPSC-derived cells were seeded into a monolayer in an animal component-free cell attachment substrate (StemCell Technologies) coated 24 well plate (ThermoScientific) and were co-cultured for 6 days with HLA-mismatched PBMCs (1:6 Target:Effector ratio) in Immunocult-XF media (StemCell Technologies) with 50 IU/ml IL-2 (StemCell Technologies). At day-0 of the experiment, the PBMCs were labeled with VPD450 proliferation dye (BD Biosciences) prior to seeding onto the monolayer. At day 6, alloreactivity of CD8⁺CD3⁺ T cell subpopulation (antibody clones SK1 and HIT3a, respectively) was assessed via proliferation (VPD450 dye dilution) and via activation, measured by the percentage of CD8⁺CD3⁺ T cells expressing CD38 (clone HIT2) and CD69 (clone FN50) through flow cytometry (CytoFlex, Beckman Colter). The dead cells were excluded with Fixable Viability Stain 780 (BD Biosciences) to determine the percentage of cells that had diluted VPD450 and thus proliferated and activated. Total CD8⁺CD3⁺ T cell and effector memory T (T_EM) cell (CD45RO ⁺ CD45RA^-CD62L^-CCR7^-) proliferation was determined by staining with the following antibodies (clones UCHL1, HI100, DREG-56, G043H7, respectively), as well as excluding dead cells with LIVE/DEAD Fixable Red stain (Thermo Fisher Scientific), to determine the percentage of cells that had diluted VPD450 and thus proliferated. Relative proliferation index was calculated by using the ratio of the percentage of proliferating cells with PSC-CM targets divided by the baseline proliferating cells without targets.

BLI in NeoThy humanized mice

NOD.Cg-KitW-41J Tyr + Prkdcscid Il2rgtm1Wjl/ThomJ (NBSGW) mice⁴⁷ (6 to 8 weeks old, male and female) were obtained from the Jackson Laboratory. NeoThy humanized mice were made⁴⁶ with neonatal cord blood and thymus tissue. Following verification of human immune cell chimerism ( > 10% human CD45⁺ and >10% human CD3⁺, by flow cytometry, as described above, using clones HI30 and UCHT1, respectively), the WT and ICAM-1 KO hPSC lines (PED05ALG at passage 47 and PED05 CD54KO ALG at passage 54, respectively), cultured as described above, were injected intramuscularly into the hind limb of the animals with Matrigel. Akaluc reporter signal was assessed via BLI using the IVIS Spectrum Imaging System (PerkinElmer) at three timepoints following the injections. AkaLumine substrate solutions were prepared dissolving 10 mg AkaLumine-HCl (AOBIOUS) in 1 mL of PBS-/- (Corning), and mice were injected intraperitoneally with 100 μL. Animals were anesthetized using isoflurane and imaged 30 min after injection. To account for variations in peak signal intensity time, images were acquired sequentially once every minute for 5 min, and the images with the highest signal were selected for analysis. Images were analyzed using Living Image 4.7 software (PerkinElmer), and maximum regions of interest were quantified in units of photons per second.

Protein collection and quantification

Cells were harvested from 6-well culture plates using TrypLE Select Enzyme (1X) (Gibco) and centrifuged at 300 g for 5 min. The cell pellet was washed twice with PBS (without Calcium or Magnesium [-/-]; Corning) buffer and resuspended in 100–200 ul of RIPA Lysis Extraction Buffer (ThermoFisher) supplemented with Phosphatase Inhibitor Mini Tablets (Thermo Scientific) and Pierce Protease Inhibitor Mini Tablets (ThermoFisher). The cell suspension was incubated on ice for 1 h to facilitate cell lysis. Mechanical disruption of cells was then performed to enhance protein extraction. The resulting cell suspension was centrifuged at 16,000 g for 20 min at 4 °C. The supernatant containing the extracted proteins was collected, snap-frozen in liquid nitrogen, and stored at −80°C for further analysis. Protein quantification was then conducted on isolated supernatant using Pierce™ Dilution-Free Rapid Gold BCA Protein Assay (ThermoFisher).

Western blot

Proteins were separated by SDS-PAGE using 4-20% Mini-PROTEAN TGX Precast Protein Gels (Bio-Rad), followed by transfer to a polyvinylidene fluoride (PVDF) membrane (MilliporeSigma). The membrane was then blocked with 5% non-fat dry milk (Kroger) in TBS-T (1X Tris Buffered Saline; Bio-Rad, with 0.5% Tween 20; Bio-Rad) for 1 h to reduce non-specific binding. After blocking, the membrane was incubated with primary antibody (β2-microglobulin [β2M] Rabbit mAb, Cell Signaling Technology; CD54/ICAM-1 Antibody, Cell Signaling Technology; GAPDH Rabbit mAb, Cell Signaling Technology) for 24 h, washed three times with TBS-T, and then incubated with secondary antibody (Anti-rabbit IgG, HRP-linked Antibody, Cell Signaling Technology) for 1 h. The membrane was washed an additional three times with TBS-T before imaging using an Azure imager. The membrane was exposed for 20–30 seconds by chemiluminescence. Molecular weights (kDa) of proteins were determined from Cell Signaling Technology data sheets (ICAM-1 = 89, 92; β2M = 12; GAPDH = 37) and verified via Precision Plus Protein Kaleidoscope Pre-stained Protein Standards (Bio-Rad).

Statistics & reproducibility

GraphPad Prism (versions 10.2.2 and 9.31) was used for statistical analysis and plotting of graphs for all experiments, except RNA sequencing, as indicated above. For leukocyte binding assays, unpaired two-sided t-tests and one-way ANOVA analysis with Tukey’s multiple comparisons test were used to determine statistical significance. For in vitro experiments, biological replicates (BR) are defined in this manuscript as independent cell culture wells treated the same throughout a given experiment, unless otherwise noted. This is in contrast to technical replicates, which we define as a repeat experimental analysis of a given BR at one specific timepoint. For in vivo studies, BR refers to an individual animal, and sex was not considered in the design of the experiments as we have not observed any sex-based differences in prior similar experiments. No statistical method was used to predetermine sample size, and no data were excluded from the analyses. The experiments were not randomized, and the investigators were not blinded to allocation during experiments and outcome assessment.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Pre-processed single cell RNA-seq gene expression data and relevant metadata are available through GEO (the Gene Expression Omnibus) under accession code GSE291617. Sanger sequencing data are available through Dryad at (https://doi.org/10.5061/dryad.xksn02vsm). Source data provided with this manuscript include adhesion assay and MLR data. Source data are provided with this paper.

Code availability

Custom code used in analyses and to produce Fig. 3d and e are available on GitHub at: https://doi.org/10.5281/zenodo.15832283.

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Acknowledgements

This study was supported in part by the Wisconsin Alumni Research Foundation (WARF), NIH NHLBI R56HL165189 (M.E.B.), NIH NIAID 75N93021C00004 (M.E.B.), NIH NHLBI U01HL134764 (T.J.K., M.E.B.), the UW Carbone Cancer Center Support Grant P30 CA014520, the UW Stem Cell and Regenerative Medicine Center Graduate Research Fellowship (S.S.), UW SciMed Graduate Research Scholars Fellowship (M.P. and L.F.), NIH NIGMS T32GM081061 (L.F.), and NIH NIDCD T32DC009401 (J.S.). The authors thank Jennifer Zellner for expert manuscript editing assistance; Andres Mejia for helpful discussions regarding teratoma pathology; Finn McCarthy for assistance with literature review; the UW Biotechnology Gene Expression Center (Research Resource Identifier— RRID:SCR_017757) for library preparation and the DNA Sequencing Facility (RRID:SCR_017759) for sequencing; Anita Bhattacharyya and Andrew Petersen at the UW Human Stem Cell Core for assistance with CRISPR/Cas9 guide RNA design and editing (supported in part by a core grant to the Waisman Center from NIH NICHD P50HD105353); Jenny Gumperz for helpful discussion and Dana Baiu for neutrophil isolation assistance; Kara R. Vogel for assistance and advice on Western Blot assays; Glen E. Leverson at the UW Department of Surgery Statistical Analysis and Research Programming Core for helpful input regarding statistical analysis; Jinhua Zhang for advice related to cardiac fibroblast differentiation; Seth Taapken and WiCell for karyotyping and quality-control reagents; and the UW Translational Research Initiative in Pathology Histology lab. Figure 2A was created with BioRender.com, via UW institutional license.

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University of Wisconsin-Madison, School of Medicine and Public Health, Department of Surgery, Madison, WI, USA
Sayandeep Saha, W. John Haynes, Jiwon Seo, Natalia M. Del Rio, Elizabeth E. Young, Alexis M. Holm, Mireya Pimentel, Liupei Huang, Wesley Blashka, Lydia Murphy, Merrick J. Scholz, Abigale Henrichs & Matthew E. Brown
Morgridge Institute for Research, Madison, WI, USA
Jue Zhang, John Steill & Ron Stewart
University of Wisconsin-Madison, School of Medicine and Public Health, Department of Medicine, Madison, WI, USA
Lauryn Flannagan, Jeremy Kratz & Timothy J. Kamp
Johns Hopkins Bloomberg School of Public Health, Baltimore, MD, USA
Jayalaxmi Suresh Babu

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Sayandeep Saha
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W. John Haynes
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Jiwon Seo
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Natalia M. Del Rio
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Elizabeth E. Young
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Jue Zhang
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Liupei Huang
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Lydia Murphy
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Jayalaxmi Suresh Babu
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John Steill
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Matthew E. Brown
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Contributions

S.S. conducted experiments, data analysis, and interpretation, and assisted in writing the manuscript. W.J.H. conducted experiments, data analysis, and interpretation, helped design experiments, provided scientific discussion, and provided helpful commentary during the writing of the manuscript. J.S. conducted experiments and data analysis. N.M.D.L. conducted experiments, data analysis, and interpretation, and assisted in writing the manuscript. E.E.Y., J.Z., A.M.H., M.P., L.F., L.H. and W.B. conducted experiments and data analysis. L.M., M.J.S., A.H. and J.S.B. conducted experiments. J.S. (Steill) analyzed data. J.K. and R.S. designed and supervised the research and analyzed data. T.J.K. designed experiments and provided critical feedback on research direction and manuscript. M.E.B. conceived the project, designed and conducted experiments, performed data analysis and interpretation, supervised the research, and wrote the manuscript.

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Correspondence to Matthew E. Brown.

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M.E.B. is a consultant for Taconic Biosciences, a founder at Allostasis, Bio, and is an inventor on a patent application filed by WARF related to this work. The other authors declare no conflict of interest.

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Nature Communications thanks Jianyi (Jay) Zhang and Ton Rabelink for their contribution to the peer review of this work. A peer review file is available.

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Saha, S., Haynes, W.J., Seo, J. et al. Diminished immune cell adhesion in hypoimmune ICAM-1 knockout human pluripotent stem cells. Nat Commun 16, 7415 (2025). https://doi.org/10.1038/s41467-025-62568-2

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Received: 10 July 2024
Accepted: 24 July 2025
Published: 12 August 2025
DOI: https://doi.org/10.1038/s41467-025-62568-2

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Abstract

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Introduction

Results

ICAM-1 blockade reduces leukocyte adhesion to hPSC-derived cellular therapies

Generation of hypoimmune ICAM-1 KO hPSCs

ICAM-1 KO hPSCs have typical differentiation capability

Surface and soluble ICAM-1 expression is ablated on differentiated cells

Immune cell adhesion is diminished in ICAM-1 KO cells

ICAM-1 KO is protective against in vitro and in vivo immune responses

ICAM-1 KO confers additional protection to first-generation hypoimmune hPSCs

Discussion

Methods

Genetic modification of PSC lines

KO of ICAM-1

Knock-In of constitutive reporter constructs

Generation of human MHC double KO-CD47 overexpression cell lines

Karyotyping

SSEA-4 flow cytometry

Alkaline phosphatase assay

PSC culture

Differentiation of cardiovascular cells from PSCs

EC differentiation

CM differentiation

cFib differentiation

Fluorescent labeling of effector immune cells

Leukocyte adhesion assays

Luminex for soluble ICAM-1

Cell surface ICAM-1 detection

Teratoma assay

HLA typing

U937 cell line

PBMC and neutrophil isolation

RNA sequencing analysis

Mixed lymphocyte reaction

BLI in NeoThy humanized mice

Protein collection and quantification

Western blot

Statistics & reproducibility

Reporting summary

Data availability

Code availability

References

Acknowledgements

Author information

Authors and Affiliations

Contributions

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Ethics declarations

Competing interests

Peer review

Peer review information

Additional information

Supplementary information

Source data

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